Over the past year, we have continued our focus on the development of new fluorescence imaging approaches. One area has been the development of a high-resolution microscopy technique capable of optical resolutions beyond the limit imposed by diffraction. This technique was developed in collaboration with Dr. Eric Betzig (Howard Hughes Medical Institute, Janelia Farm Research Campus and New Millennium Research), Dr. Harald Hess (HHMI, Janelia Farm and NuQuest Research) and members of the Juan Bonifacino lab (CBMB), and the Michael Davidson lab (Florida State University). Termed photoactivated localization microscopy (PALM), the method involves serial photoactivation and subsequent bleaching of numerous sparse subsets of photoactivated fluorescent protein molecules. Individual molecules are localized at near molecular resolution by determining their centers of fluorescent emission via a statistical fit of their point-spread-function. The aggregate position information from all subsets is then assembled into a super-resolution image, in which individual fluorescent molecules are isolated at high molecular densities (up to 105 molecules/m2). PALM imaging of intracellular structures (including lysosome, Golgi apparatus and mitochondria) in cryo-prepared thin sections was demonstrated, as well as imaging of vinculin and actin in fixed cells with TIRF excitation, and correlative PALM/transmission electron microscopy of a mitochondrial marker protein. We have also worked toward developing dual-label PALM using two different photactivatable molecules expressed within cells. In addition, we have developed a system for doing single particle tracking using PALM in living cells that allows protein diffusion and immobilization to be characterized at the single molecule level.[unreadable] A second new fluorescent protein technique developed in our lab allows a proteins topology to be determined in living cells. Termed fluorescence protease protection (FPP), the assay provides a fluorescent readout in response to trypsin-induced destruction of GFP attached to a protein-of-interest before and after plasma membrane permeabilization. In performing the FPP assay, a fluorescent protein is attached to the N or C terminus of a protein of interest. Subsequently, cells expressing the fusion protein are exposed to trypsin either before or after plasma membrane permeabilization by digitonin. If the fluourescent protein moiety on the expressed protein faces the environment exposed to trypsin (that is the cytoplasm), then its fluorescent signal will be lost. Conversely, if the fluorescent protein moiety on the expressed protein faces the environment protected from trypsin (that is, the lumen of a compartment) then its fluorescence persists. Given these outcomes and the fluorescent proteins known engineered position within the protein, it is possible to deduce the orientation of the protein across the lipid bilayer. We demonstrated the broad applicability of FPP by using it to define the topology of proteins localized to several different organelles, including the ER, Golgi apparatus, mitochondria, peroxisomes and autophagosomes. [unreadable] A third new fluorescent labeling technique we developed was in cellulo pulse chase analysis. In this approach, a small area of the cell expressing a photoactivatable protein is 'switched on' using an activating pulse of light. The fate of the newly fluorescent proteins in this area are then followed over time. Since only the molecules that have been switched on during the photoactivation event are fluorescent, it becomes possible to follow the fate of these molecules without concern that new molecules will become fluorescent during the subsequent imaging period. We used this photo-labeling, pulse-chase strategy for distinguishing newly synthesized from previously synthesized peroxisomal protein components and for visualizing both old and new peroxisomes within cells. Peroxisomes are small-membrane bound organelles that function in many metabolic processes, including metabolism of fatty acids and conversion of perioxides to nontoxic forms. We found that old peroxisomes contained both newly synthesized and previously synthesized protein components, whereas new peroxisomes contained only newly synthesized peroxisomal protein components. This argued against fission being the predominant mechanism for mammalian peroxisome formation and indicated that de novo biogenesis of peroxisomes from the ER was important for maintenance of peroxisomes under normal conditions.